Protein therapeutics is the fastest growing segment of the biotechnology and pharmaceutical industry. Protein therapeutics includes monoclonal antibodies, recombinant proteins, chimeric proteins and other protein receptor constructs. This segment is expected to reach over $70 billion in sales by 2011.
A major hurdle in the development and use of proteins as pharmaceutical drugs is the ability to store, transport and deliver them in a safe stable form. It is well known that factors, such as temperature, solvent, ligands, excipients, pH, and salt concentration, affect a particular protein's stability. The identification of buffer, ligand and excipient conditions that maximize the stability and eliminate protein aggregation is critical during development and often requires the evaluation of hundreds of conditions. This combination of buffer, ligand and excipients conditions is referred to as the storage formulation throughout this disclosure. Unfortunately, it is difficult to vary all of the various parameters to determine the ideal storage formulation for a particular protein.
There are different ways to measure protein stability and each involves disrupting the protein structure through either physical or chemical means. This disruption of the protein structure is referred to as denaturation.
Temperature is one of the most widely used physical denaturants. In this scenario, a protein is subjected to increasing temperature and the corresponding changes in its structure are recorded. One of the disadvantages of temperature denaturation is that proteins typically denature at temperatures at or above 60° C. However, in most instances, the temperatures of interest are physiological (about 37° C.), room (about 25° C.) and storage (4° C.). Thus, results from temperature-based denaturation tests must be extrapolated by more than 25° C. to understand the effects at the temperatures of interest. In addition, most proteins used as biologics undergo irreversible temperature denaturation, which precludes a meaningful calculation of thermodynamic stability at the temperatures of interest. In addition, a formulation that elicits a higher denaturation temperature does not necessarily result in a more stable protein at room temperature.
A second way to measure protein stability is through the use of chemical denaturants, such as urea or guanidine hydrochloride. This method permits measurements to be done at any desired temperature.
The structural stability of a protein is determined by its Gibbs energy of stability, ΔG. This value, ΔG, is a function of temperature, chemical denaturants and other physical and chemical variables. Using the common example of a two state model, where a protein is either folded (i.e. native) or unfolded (i.e. denatured), the protein can transition between these two states:                NU, wherein N is the native (folded) state and U is the unfolded state.        
Two different rate constants can be defined from this transitional equation. Kf is the rate of the folding reaction; while Ku is the rate of the unfolding reaction. Finally, the equilibrium constant, K, can be defined as the ratio of the unfolding rate to the folding rate, or
  K  =                    K        u                    K        f              .  Furthermore, the Gibbs energy can be expressed in terms of K, asΔG=−RT ln(K),where R is the gas constant, T is the temperature, expressed in Kelvin and ln(K) is the natural log of K. Thus, if K is greater than one, the protein unfolds at a higher rate than it folds, and its Gibbs energy is negative. Conversely, if K is less than one, the protein unfolds at a slower rate than it folds, and its Gibbs energy is positive. Also, K is equal to the ratio of the concentration of protein in the unfolded state and the concentration of protein in the folded state K=[U]/[F].
In addition, it has been observed that, for chemical denaturants, a nearly linear relationship exists between the Gibbs energy and the concentration of the denaturant. This relationship may be expressed asΔG=ΔG0−m*[denaturant],where ΔG0 is the intrinsic Gibbs energy, [denaturant] is the concentration of denaturant, and m is the multiplier, which is unique for a particular protein.
For a native/unfolded equilibrium, the fraction of protein molecules which are unfolded, or denatured, Fd, is given by:
            F      d        =          K              1        +        K              ,where K is the equilibrium constant.
This equation can be used to allow calculation of a denaturation curve. When a protein changes from its folded state to an unfolded state, certain measurable characteristics of the protein also change. One such characteristic is the fluorescence of the protein.
While the preferred embodiment described in this application utilizes fluorescence emission (intrinsic or extrinsic) as a way to determine the degree of denaturation or unfolding of a protein, the disclosure is not limited to this technique. There are many physical observable properties and their associated instrumentation, in addition to fluorescence spectroscopy, that are sensitive to the degree of denaturation of a protein. These observable properties include, but are not limited to uv/vis spectroscopy, circular dichroism, nuclear magnetic resonance (NMR), infrared spectroscopy (IR) among others.
FIG. 1 shows a typical urea denaturation curve for an antibody. The y, or vertical, axis is a measure of the intrinsic fluorescence of the protein. The fluorescence of different dyes, usually known as protein probes, can also be used. The horizontal, or x, axis is the concentration of urea in solution with the protein. As can be seen, at a certain point, between 3M and 4M urea, the fluorescence of the protein changes dramatically, indicating that it has denatured.
The generation of the data needed to produce such a graph is laborious. In one scenario, a solution containing the protein and any excipients is prepared. A sample of this solution is then subjected to fluorescent light and the emission is recorded. This is the baseline fluorescence with no chemical denaturant. In some embodiments, an amount of urea is then added to the remainder of the solution, and the light test is repeated on a portion of this modified solution. An additional amount of urea is then added to the remainder of the solution and a third light test is performed. This process is repeated for the number of desired samples. The amount of urea added each time is a function of the desired granularity of the test, and the range of urea molarities to be included. Such a method is prone to errors, as there are cumulative errors due to the constant addition of urea to the remaining solution. In this stepwise urea addition method, the process will result in the dilution of the protein and also a smaller fluorescence signal. In addition, since the solubility of urea is about 10.5M and a final 8M urea concentration is needed, the starting protein solution volume needs to be extremely small. The protein will be significantly diluted as the experiment progresses.
In another embodiment, a plurality of solutions, each with the protein, any excipients, and the proper amount of urea, is individually prepared. Each of these prepared solutions is then light tested to determine its fluorescence. While this method removes the cumulative errors associated with the previous method, it is extremely time consuming, especially for a large number of samples.
The resulting graph, such as that shown in FIG. 1, shows the stability of a particular combination of buffer, ligand and excipient conditions in the presence of a chemical denaturant. More stable combinations have a similarly shaped graph, shifted to the right. Conversely, less stable combinations have a graph shifted to the left. The goal of this testing is to find a combination with the maximum stability in the presence of the chemical denaturant. This combination can then be used as the storage formulation for the protein as it is stored and shipped.
Given the increased importance of developing proteins for pharmaceutical purposes, there is a dearth of systems and methods available to aid in the determination of the ideal storage formulation in which the protein is most stable.
For example, denaturation graphs are an effective way to understand the stability of a protein in a particular buffer solution. However, as described above, the creation of denaturation graphs is tedious and error prone. Furthermore, the testing required to fully understand the effect of changing one or more components of that buffer solution is so labor intensive that it is rarely performed. A system and method for automatically creating a plurality of denaturation graphs would be beneficial.